JBC Transcription and Nuclear Factor Monoclonals

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Originally published In Press as doi:10.1074/jbc.M103894200 on August 30, 2001

J. Biol. Chem., Vol. 276, Issue 47, 43516-43523, November 23, 2001
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Prion Protein Fragment PrP-(106-126) Induces Apoptosis via Mitochondrial Disruption in Human Neuronal SH-SY5Y Cells*

Conor N. O'Donovan, Deirdre Tobin, and Thomas G. CotterDagger

From the Tumor Biology Laboratory, Biochemistry Department, University College Cork, Lee Maltings, Prospect Row, Cork, Ireland

Received for publication, May 1, 2001, and in revised form, August 22, 2001


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

The synthetic peptide PrP-(106-126) has previously been shown to be neurotoxic. Here, for the first time, we report that it induces apoptosis in the human neuroblastoma cell line SH-SY5Y. The earliest detectable apoptotic event in this system is the rapid depolarization of mitochondrial membranes, occurring immediately upon treatment of cells with PrP-(106-126). Subsequent to this, cytochrome c release and caspase activation were observed. Caspase inhibitors demonstrated that while the peptide activates caspases they are not an absolute requirement for apoptosis. Parallel to caspase activation, PrP-(106-126) was also observed to trigger a rise in intracellular calcium through release of mitochondrial calcium stores. This leads to the activation of calpains, another family of proteases. A calpain inhibitor demonstrated that while calpains are activated by the peptide they also are not an absolute requirement for apoptosis. Interestingly a combination of caspase and calpain inhibitors significantly inhibited apoptosis. This illustrates alternative pathways leading to apoptosis via caspases and calpains and that blocking both pathways is required to inhibit apoptosis. These results implicate the mitochondrion as a primary site of action of PrP-(106-126).


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Prion-related encephalopathies are a family of neurodegenerative disorders including conditions such as scrapie in sheep, bovine spongiform encephalopathy in cattle, and Creutzfeldt-Jakob disease and Gerstmann-Straussler-Scheinker syndrome among others in humans. They are characterized by vacuolation of the neuropil, neuronal loss, and gliosis (1). In many cases this is also accompanied by the extracellular accumulation of the scrapie isoform (PrPSC)1 of the normal cellular prion protein (PrPC), which can aggregate into fibrils in the extracellular matrix (2).

PrPSC is widely believed to be the infectious agent of these diseases (3), and the formation of PrPSC is thought to be via a post-translational conformational change by which PrPC complexes with PrPSC to yield two molecules of PrPSC (4). PrPC is a cell surface protein mainly expressed in the neuronal and glial cells of the central nervous system. The exact function of PrPC remains unknown, although recent studies have implicated it in copper metabolism (5, 6) and signal transduction (7). Its expression is necessary for the pathogenesis of the spongiform encephalopathies (8).

Apoptosis is a physiologically important cellular suicide pathway, which has also been implicated in a number of pathological conditions (9). There is some evidence to indicate that the mechanism of neuronal cell death in prion diseases is apoptosis as apoptotic neurons have been observed in the brain of scrapie-infected sheep (10), the brain and retinae of mice infected with the 79A strain of scrapie (11), and the brain of human Creutzfeldt-Jakob disease patients (12).

A synthetic peptide corresponding to residues 106-126 of human PrP (PrP-(106-126)) has previously been found to induce apoptosis in primary rat hippocampal cultures (13), primary mouse cerebellar cultures (14), the rat pituitary clonal cell line GH3 (15), and more recently in mouse retinae in vivo (16). PrP-(106-126) is one of a number of peptides corresponding to sequences within fragments of amyloid proteins isolated from the brains of patients suffering from Gerstmann-Straussler-Scheinker syndrome (17). It retains the ability of PrPSC to aggregate into amyloid-like fibrils and the tendency to adopt a mostly beta -sheet structure (18). Residues 106-126 of PrP constitute a region maintained in all the PrP isoforms that have been found to accumulate in the brains of patients suffering from prion diseases. Under normal physiological conditions a catabolic pathway also leads to cleavage of this region of the prion protein at residues 110 and 111 (19).

The importance of the 106-126 sequence of the prion protein makes it a useful model for the in vitro study of prion-induced cell death. In this study we determine, for the first time, the effect of PrP-(106-126) in human neuroblastoma cells in vitro and demonstrate its ability to induce apoptosis. Furthermore, we establish the mitochondrion as a target of PrP-(106-126) action and show that the peptide activates two distinct biochemical pathways involving caspases and calpains, which both result in apoptosis.

    EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Peptides and Drugs-- PrP-(106-126) (KTNMKHMAGAAAAGAVVGGLG) and PrP-(106-126) scrambled (scr) (AVHTGLGAMAALNMVVGGAAGL) were synthesized and purified by MWG Biotech (Milton Keynes, UK). Peptides were dissolved in sterile phosphate-buffered saline (PBS) to a concentration of 1 mM before use and were freshly prepared before each experiment. The inhibitors z-VAD-fmk (Enzymes Systems Products, Livermore, CA) and calpeptin (Sigma) were added to cells 15 min prior to drug/peptide treatments.

Thioflavin-T Binding Assay-- The degree of peptide aggregation was measured fluorimetrically by thioflavin-T binding (20). Aged preparations of peptides in PBS were added to 50 mM glycine, pH 9, with 2 µM thioflavin-T (Sigma) to a concentration of 100 µM. Samples were incubated at room temperature for 5 min, and then fluorescence was measured on a Spectramax Gemini fluorometer with excitation and emission maxima of 435 and 485 nm, respectively. Samples were prepared in triplicate.

Cell Culture-- The adherent human neuroblastoma cell line SH-SY5Y was cultured in RPMI 1640 medium (Life Technologies, Inc.) supplemented with 10% fetal calf serum (Life Technologies, Inc.), 1% penicillin/streptomycin (Life Technologies, Inc.), and 1% L-glutamine (Life Technologies, Inc.). Cells were maintained at 37 °C in a humidified 5% CO2 atmosphere. For experiments, cells were maintained in serum-free RPMI 1640 medium supplemented with 1% Growth Medium Supplement-A (Life Technologies, Inc.), 1% penicillin/streptomycin, and 1% L-glutamine. When being passaged or harvested for analysis cells were lifted using trypsin/EDTA.

MTT Assay-- Cytotoxicity was assessed by the conversion of MTT (Sigma) to a formazan product. After appropriate incubation of cells with peptides, MTT was added to each well to a final concentration of 0.25 mg/ml and then incubated for 4 h at 37 °C. Microtiter plates were then centrifuged at 200 × g for 5 min. The reaction was terminated by removal of the supernatant and addition of 100 µl of Me2SO to each well. Following thorough mixing to dissolve the formazan product, the plates were read at 620 nm on a microELISA plate reader. Assays were performed in replicate of four samples.

Annexin V Binding-- Cell viability was assessed by flow cytometry that monitored annexin V binding and propidium iodide (PI) uptake simultaneously. After appropriate incubation with drugs/peptides, cells were resuspended in annexin V binding buffer and then treated with annexin V (1×) and PI (5 µg/ml) for 5 min at room temperature. Samples were then analyzed by fluorescence on a FACScan flow cytometer (Becton Dickinson, Oxford, UK). Fluorescence was measured through a 530/30 band pass filter (FL-1) to monitor annexin V binding and through a 585/42 band pass filter (FL-2) to monitor PI uptake. An initial increase in FL-1 fluorescence is indicative of apoptosis before an increase in FL-2, indicative of secondary necrosis, is observed.

DNA Fragmentation Assay-- DNA was isolated and electrophoresed on 10% agarose gels by the method of McGahon et al. (21).

Western Blotting-- Cells were lysed in RIPA buffer (50 mM Tris-HCl (pH 7.4), 1% Nonidet P-40, 0.25% sodium deoxycholate, 1 mM EGTA, 1 mM Na3VO4, 1 mM NaF, 150 mM NaCl, 0.1 mM phenylmethylsulfonyl fluoride, and 1 µl of protein inhibitor mixture (1 µg/ml antipain, 1 µg/ml aprotinin, 1 µg/ml chymostatin, 0.1 µg/ml leupeptin, and 1 µg/ml pepstatin)) and put on ice for 40 min. Samples were then centrifuged at 20,800 × g for 10 min. The supernatant was then transferred into Eppendorf tubes, and protein concentrations were determined using a Bio-Rad protein assay reagent. Protein was separated by SDS-polyacrylamide gel electrophoresis (15% gel, 30 µg of protein/sample) and then transferred to a nitrocellulose membrane. Proteins were detected using appropriate antibodies and the ECL detection reagent (Amersham Pharmacia Biotech). The antibodies used were mouse monoclonal anti-human Bcl-2 (DAKO, Cambridge, UK), rabbit monoclonal anti-Bcl-XL, (Calbiochem), rabbit monoclonal anti-Bax (DAKO, Cambridge, UK), monoclonal anti-cytochrome c (DAKO), and monoclonal anti-cytochrome c oxidase (DAKO).

Measurement of Mitochondrial Membrane Potential (Psi m)-- Cells were harvested and treated with 10 µM JC-1 (Molecular Probes, Leiden, Netherlands) for 15 min at 37 °C (22). Mitochondrial membrane potential was then measured by fluorescence emission on a FACScan flow cytometer (Becton Dickinson). Fluorescence emission was collected through FL-1 on a log scale. FL-1 measures the fluorescence of JC-1 monomers, which increase in number as the mitochondrial membrane depolarizes.

Detection of Caspase-3 Activity-- Cells were washed in 2 ml of PBS, fixed in 1% paraformaldehyde on ice for 30 min, and then washed in 1 ml of IFA-Tx (4% fetal calf serum, 10 mM HEPES (pH 7.4), 0.1% sodium azide, 0.1% Triton X-100, 150 mM saline). Samples were then resuspended in 150 µl of IFA-Tx containing primary antibody (rabbit polyclonal anti-active caspase-3, 1:600 dilution (Pharmingen, San Diego, CA)), put on ice for 1 h, centrifuged at 200 × g, and washed twice in 1 ml of IFA-Tx. IFA-Tx was aspirated off, and to the remaining drop in each sample, 2 µl of secondary antibody (fluorescein isothiocyanate-labeled anti-rabbit (Sigma)) was added. Samples were left for 1 h on ice in darkness and then washed twice in 1 ml of IFA-Tx. Samples were then resuspended in 0.5 ml of PBS, and fluorescence was measured on a FACScan flow cytometer through FL-1. An increase in FL-1 fluorescence above the control is indicative of caspase activity. Samples treated with an irrelevant antibody (rabbit IgG (Sigma)) and with secondary antibody only were used as controls.

Calpain Activity Assay-- Cells were washed once in PBS and then resuspended and lysed in 40 µl of 10 mM HEPES, 50 mM NaCl on ice for 40 min. After lysis the samples were centrifuged at 1000 × g for 5 min to pellet the membrane fraction. The supernatant (cytosolic fraction) was transferred to a clean Eppendorf tube, and the membrane fraction was resuspended in 40 µl of 10 mM HEPES, 50 mM NaCl. A protein determination assay was performed on both fractions, and an equal amount of protein was loaded into each well of a 96-well plate (ideally 100 µg of protein/well). To each well, 32 µl of fluorescent substrate (Suc-Leu-Tyr (AFL-117) (Enzyme Systems Products)) was added, and the total volume of each well was brought to 200 µl with imidazole buffer (100 mM imidazole, 5 mM L-cysteine, 1 mM mercaptoethanol, 10 mM CaCl2, 4% Me2SO in H2O). Samples were incubated at 37 °C for 30 min after the addition of the substrate. Fluorescence was then measured on a fluorometer (Spectra Max Gemini, Molecular Devices) with excitation and emission wavelengths of 400 and 505 nm, respectively.

Glutathione Assay-- Cells were washed once in PBS and then resuspended in 650 µl of PBS. Next 3.25 µl of 10 mM monochlorobimane (Molecular Probes) was added to each sample. Samples were measured in triplicate, so 200 µl of each sample was aliquoted into three wells of a dark 96-well plate. Samples were incubated at room temperature for 15 min in darkness and then read on a fluorometer (Spectra Max Gemini, Molecular Devices) with excitation and emission maxima of 395 and 482 nm, respectively.

Catalase Assay-- The activity of catalase was measured by following the decrease in absorbance at 240 nm due to H2O2 decomposition (23). Activity was measured as rate of change of absorbance (Delta O.D./min).

Calcium Measurements-- Changes in intracellular calcium concentration were detected by loading cells with FLUO-3 (1×) (Molecular Probes) for 15 min prior to sample collection and measurement of fluorescence on a FACScan flow cytometer. Fluorescence emission was collected through FL-1 on a log scale, and an increase in FL-1 fluorescence is indicative of an increase in calcium levels.

Isolation of Mitochondria and Cytochrome c Detection-- Cells were harvested, washed in 1 ml of homogenizing medium (1 M sucrose, 1 M Tris-HCl (pH 7.4), 50 mM EGTA, and 1% bovine serum albumin), resuspended in 80 µl of homogenizing medium, and Dounce-homogenized. Samples were centrifuged at 1000 × g for 5 min at 4 °C, and the supernatant was transferred to a clean Eppendorf tube. The supernatant was centrifuged at 10,000 × g for 5 min at 4 °C. The pellet was then washed in 100 µl of Wash 1 (1 M sucrose, 1 M Tris-HCl, 1% bovine serum albumin (pH 7.4), 50 mM EGTA, and 1 M KCl), centrifuged at 10,000 × g for 5 min at 4 °C, and washed in 100 µl of Wash 2 (1 M mannitol, 1 M KCl, 1 M Tris-HCl, 1% bovine serum albumin (pH 7.4)). Finally the sample was centrifuged at 10,000 × g for 5 min at 4 °C and resuspended in RIPA buffer (see "Western Blotting"). Cytochrome c was visualized by Western blot analysis using anti-cytochrome c (Pharmingen). The supernatant was retained as the cytosolic fraction.

    RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

PrP-(106-126) Forms Aggregates and Induces Apoptosis in SH-SY5Y Cells-- PrP-(106-126) has previously been reported to induce cell death as a result of its ability to form aggregates (13, 24). The aggregation status of the peptide in this study was ascertained by means of a thioflavin-T assay (Fig. 1A). Thioflavin-T fluorescence increases in the presence of protein aggregates. PrP-(106-126) was seen to aggregate immediately at day 0 when prepared at 100 µM in PBS, and over time it gradually increases its aggregation status. The scrambled version of the peptide shows little aggregation even over time. PrP-(106-126) was observed to induce cell death in a dose-dependent manner over time as measured by MTT assay (Fig. 1B). MTT is converted to a formazan product by mitochondrial enzymes, which become inactive as the cell dies. Measurement of this formazan product is an indicator of cell viability. The mechanism of cell death induced by PrP-(106-126) was shown to be apoptosis by annexin V binding (Fig. 1C). Annexin V binds to phosphatidylserine, which flips from the inner to the outer leaflet of the cell membrane during apoptosis (73). Apoptosis could be detected as early as 2 h after treatment when 22% of the cell population were annexin-positive. Cells were incubated with annexin V and PI simultaneously The population of cells in the lower left quadrant represents viable cells (Fig. 1C). An increase in FL-1 represents annexin V-positive cells in the lower right quadrant. This is the apoptotic population. The final shift in FL-2 up to the top right quadrant represents PI-positive cells, indicative of membrane permeability and secondary necrosis. The percentage of the overall population in each quadrant is given in the circles. The mechanism of cell death was further confirmed to be via apoptosis by the observation of DNA ladder patterns upon electrophoresis of isolated DNA from peptide treated cells (Fig. 1D). The internucleosomal cleavage of DNA into ~200-base pair fragments is a typical biochemical hallmark of apoptosis in a number of systems (74). No effects were observed upon treatment with PrP-(106-126)scr.


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  		Fig. 1.   Peptide aggregation status and its induction of apoptosis in the SH-SY5Y cell line. A, PrP-(106-126) was found to aggregate immediately when prepared in PBS at a concentration of 100 µM as measured by Thioflavin-T fluorescence. Its aggregation was observed to increase slowly over 8 days. PrP-(106-126)scr showed a much lesser degree of aggregation that remains constant over time. Results are expressed as the mean ± S.E. B, PrP-(106-126) (black-square) was seen to induce death in the SH-SH5Y cell line in a dose-dependent manner as measured by the MTT assay at 24 h. PrP-(106-126)scr (black-diamond ) induced no death. Results are expressed as the mean ± S.E. C, 100 µM PrP-(106-126) induces apoptosis within 2 h as compared with controls, which were untreated or treated with 100 µM PrP-(106-126)scr. Apoptosis was measured by annexin V binding via flow cytometry (represented by an increase in FL-1). Secondary necrosis is indicated by a subsequent increase in FL-2. The percentage of the overall population in each quadrant is given in the circles. D, DNA ladder patterns confirm that death induced by PrP-(106-126) is apoptotic in nature. Lane A, untreated; lane B, 0.5 µM staurosporine as positive control; lane C, 100 µM PrP-(106-126), 24 h; lane D, 100 µM PrP-(106-126), 48 h; lane E, 100 µM PrP-(106-126)scr.

Rapid Intracellular Apoptotic Events in Response to PrP-(106-126)-- A number of typical apoptotic events are observed to occur after treatment of cells with 100 µM PrP-(106-126). Mitochondrial dysfunction characterized by a loss of transmembrane potential has been found to be a central event in many cases of apoptosis (25). Analysis of Psi m using the lipophilic fluorescent probe JC-1 demonstrates extensive loss of Psi m (represented by an increase in fluorescence of FL-1 due to increased numbers of JC-1 monomers) within 15 min of treatment with 100 µM PrP-(106-126) (Fig. 2A). PrP-(106-126)scr (100 µM) had no significant effect. Although the response is maximal at 15 min, mitochondrial membrane depolarization is observed to commence immediately upon treatment of the cells with PrP-(106-126) when fluorescence in FL-1 is measured as a function of time on the FACScan flow cytometer (Fig. 2B). Also within this 15-min time frame, cytochrome c is released from the mitochondria into the cytosol (Fig. 2C), and caspase-3 activity is easily detectable by measuring the fluorescence of an anti-active caspase-3 antibody on a flow cytometer (Fig. 2D). Cytochrome c is a protein of the mitochondrial intermembrane space that is commonly released during apoptosis following membrane depolarization, which leads to caspase activation. The caspases are a family of serine threonine proteases responsible for the cleavage of a number of substrates during apoptosis (42). Despite these early apoptotic events, at 15 min apoptosis is not detectable by annexin binding.


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  		Fig. 2.   Mitochondrial depolarization, caspase activation, and cytochrome c release are all early events in apoptosis induced by PrP-(106-126). A, extensive depolarization of the mitochondrial membrane represented by an increase in FL-1 was observed within 15 min of treatment with 100 µM PrP-(106-126) using the fluorescent probe JC-1. · · ·, 100 µM PrP-(106-126); ---, untreated control; - - -, 100 µM PrP-(106-126)scr. B, a time course of this event measured by flow cytometry, again using JC-1, revealed that depolarization begins immediately upon treatment with the peptide. C, within the same time frame cytochrome c is released from the mitochondria as measured by Western blot analysis. Lane 1, untreated control; lane 2, 100 µM PrP-(106-126); lane 3, 100 µM PrP-(106-126)scr. Controls of untreated cells show that cytochrome c (Cyt-c) resides in the mitochondrial fraction, and cytochrome c oxidase (an integral mitochondrial membrane protein) is used as a marker for the purity of the cytosolic (C) and mitochondrial (M) fractions. D, caspase-3 activity, as measured by an increase in fluorescence of an anti-active caspase-3 antibody on a flow cytometer, is also observed within 15 min of peptide treatment. - - -, 100 µM PrP-(106-126)scr; ---, untreated control; ---, 100 µM PrP-(106-126).


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  		Fig. 3.   Caspases target antiapoptotic Bcl-2 yet are not necessary for apoptosis induced by PrP-(106-126). A, the levels of a number of Bcl-2 family members were examined by Western blot analysis. Bcl-2 was observed to be degraded within 15 min of treatment with 100 µM PrP-(106-126). Other Bcl-2 family members, including Bcl-XL and Bax, were not affected. Lane 1, untreated; lane 2, 100 µM PrP-(106-126); lane 3, 100 µM PrP-(106-126)scr. B, this degradation of Bcl-2 was observed to be caspase-dependent as it could be blocked by z-VAD, a pan caspase inhibitor. Lane 1, untreated; lane 2, 100 µM z-VAD; lane 3, 100 µM PrP-(106-126); lane 4, 100 µM PrP-(106-126) + 100 µM z-VAD; lane 5, 100 µM PrP-(106-126) + 200 µM z-VAD; lane 6, 100 µM PrP-(106-126)scr. C, Z-VAD blocks caspase-3 activity as measured by flow cytometry. ---, untreated control; · · ·, 100 µM PrP-(106-126)scr; ---, 100 µM PrP-(106-126). D, blocking caspase activity and Bcl-2 breakdown does not inhibit mitochondrial depolarization. ---, untreated control; · · ·, 100 µM PrP-(106-126)scr; - - -, 100 µM PrP-(106-126) + 100 µM z-VAD; ---, 100 µM PrP-(106-126).

Bcl-2 Protein Levels Are Reduced in Response to PrP-(106-126)-- In view of the rapid effects of PrP-(106-126) on the mitochondria we investigated possible effects on the Bcl-2 family of proteins. These are important regulators of apoptosis, composed of both pro- and antiapoptotic members, and are known to operate at the level of the mitochondrion (26-28). We examined the intracellular levels of Bcl-2, an antiapoptotic protein, when cells were treated with 100 µM PrP-(106-126) (Fig. 3A). After a 15-min treatment with 100 µM PrP-(106-126) there was a dramatic reduction of Bcl-2 protein as compared with the untreated control. PrP-(106-126)scr (100 µM) caused no change in the levels of Bcl-2. We also examined the effects of the peptide on other Bcl-2 family members, Bcl-XL (antiapoptotic) and Bax (proapoptotic) (Fig. 3A). In contrast to what was observed with Bcl-2, we found no change in the intracellular levels of Bcl-XL or Bax in response to 100 µM PrP-(106-126). As a caspase cleavage site was previously reported to be present in Bcl-2 (29), the effect of z-VAD, a pan caspase inhibitor previously reported to block apoptosis, on Bcl-2 degradation was investigated (Fig. 3B). The reduction of Bcl-2 induced by 100 µM PrP-(106-126) was found to be inhibited in cells that were pretreated with 100 or 200 µM z-VAD. Cells treated with 200 µM z-VAD or 100 µM PrP-(106-126)scr alone showed no change in their Bcl-2 levels. A concentration of 100 µM z-VAD was shown to block caspase-3 activity as measured by anti-active caspase-3 antibody on the FACScan flow cytometer (Fig. 3C). Thus, the inhibition of caspases maintains the protein levels of the antiapoptotic Bcl-2 even in the presence of PrP-(106-126). The same treatment of cells with z-VAD did not inhibit mitochondrial depolarization (Fig. 3D), placing this event upstream of caspase activation and Bcl-2 degradation. More importantly, z-VAD did not inhibit apoptosis despite blocking caspase-mediated Bcl-2 degradation, indicating that caspases are not necessary for PrP-(106-126) to induce cell death.

Oxidative Stress Was Not Induced by PrP-(106-126)-- With caspases found to be nonessential for apoptosis in this system, another mechanism must be at work. Maintaining focus on the mitochondrion, as this was the site of the earliest observed effect of PrP-(106-126), we investigated the possible role of oxidative stress due to disruption of mitochondrial function as this is the primary site of intracellular reactive oxygen species production. Oxidative stress has been shown to have a role in a number of apoptotic systems (30, 31). We examined the intracellular levels of peroxides, superoxide anion, and nitric oxide as well as the activities of Mn-superoxide dismutase and Cu,Zn-superoxide dismutase (data not shown). We also looked at glutathione levels using the fluorescent monochlorobimane probe (Fig. 4A) and the activity of catalase by following the decrease in absorbance at 240 nm due to H2O2 decomposition (Fig. 4B) in response to PrP-(106-126). Glutathione not only acts as a reactive oxygen species scavenger but also functions in the regulation of the intracellular redox state, and catalase is the primary defense mechanism against H2O2. None of the aforementioned changed appreciably in response to PrP-(106-126). Staurosporine (0.25 µM) and valinomycin (50 nM) were used as positive controls as both these drugs induce oxidative stress in SH-SY5Y cells. The peptide was not found to predispose the cells to death by a secondary oxidative insult either (data not shown).


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  		Fig. 4.   Oxidative stress is not involved in apoptosis induced by PrP-(106-126). A, GSH levels were not observed to change in response to 100 µM PrP-(106-126). Buthionine sulfoximine, a known depletor of GSH, was used as a positive control for GSH depletion, and the drugs staurosporine and valinomycin, known to induce oxidative stress, were used as positive controls. Column 1, untreated control; column 2, 100 µM PrP-(106-126); column 3, 100 µM PrP-(106-126)scr; column 4, 0.25 µM staurosporine; column 5, 50 nM valinomycin; column 6, 1 mM buthionine sulfoximine. B, no changes in the activity of catalase, as measured by the decrease in absorbance at 240 nm of H2O2, were observed in response to 100 µM PrP-(106-126). Valinomycin and staurosporine were again used as controls. Column 1, untreated control; column 2, 100 µM PrP-(106-126); column 3, 100 µM PrP-(106-126)scr; column 4, 0.25 µM staurosporine; column 5, 50 nM valinomycin. §, p < 0.05.

Calcium Homeostasis Was Altered by PrP-(106-126)-- Another function of the mitochondrion is in the regulation of intracellular calcium levels, therefore we investigated whether PrP-(106-126) could be exerting an effect through the deregulation of calcium homeostasis. Using the fluorescent probe FLUO-3 we demonstrated an intracellular rise in calcium levels (represented by an increase in FL-1 fluorescence) in response to PrP-(106-126) (Fig. 5A). The response is immediate, and calcium levels reach a sustained peak by 30 s. This experiment was repeated in the presence of EGTA, a calcium chelator, to show that the source of the calcium was intracellular (data not shown). BAPTA-AM, an intracellular calcium chelator, was found to inhibit this increase in calcium levels, confirming the intracellular nature of the source of the calcium rise (Fig. 5B). The two major calcium stores in the cell are the endoplasmic reticulum and the mitochondria. To determine which was releasing calcium into the cytosol in this system we used thapsigargin, which causes a rapid release of endoplasmic reticulum calcium stores. Upon treatment of cells with thapsigargin, an increase in calcium levels was observed using the FLUO-3 probe. Once that response had reached its maximum, cells were treated with 100 µM PrP-(106-126), and a further increase in calcium was observed (Fig. 5C). This indicates that PrP-(106-126) releases calcium from a site other than the endoplasmic reticulum. The mitochondria make up the only other store of calcium in the cell large enough to account for the PrP-(106-126)-induced rise in calcium levels.


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  		Fig. 5.   PrP-(106-126) triggers a rapid rise in intracellular calcium, which is not from an extracellular or an endoplasmic reticular source. A, an immediate increase in intracellular calcium was observed using the fluorescent probe FLUO-3 upon treatment with 100 µM PrP-(106-126). Maximal calcium levels were reached in 30 s. ---, untreated control; ---, 100 µM PrP-(106-126). B, the intracellular calcium chelator BAPTA-AM (1 mM) was observed to block this increase in calcium levels. ---, untreated control; · · ·, 100 µM PrP-(106-126) + 10 µM BAPTA-AM; ---, 100 µM PrP-(106-126). C, using thapsigargin to release endoplasmic reticular stores of calcium before treatment with PrP-(106-126), a further increase in calcium was still observed upon treatment with PrP-(106-126), indicating a non-endoplasmic reticulum source for the PrP-(106-126)-induced increase in calcium. ---, untreated control; · · ·, 100 µM PrP-(106-126); ---, 1 µM thapsigargin; - - -, 1 µM thapsigargin + 100 µM PrP-(106-126).

Calpain Activity Increased in Response to PrP-(106-126)-- We next examined the activity of calpains in response to the PrP-(106-126)-induced rise in intracellular calcium. The calpains are a group of calcium-activated proteases that have also been implicated in apoptosis. They can exist in active and inactive forms associated with the cell membrane or the cytosol (32-34). Within 15 min of treating cells with 100 µM PrP-(106-126) an increase in calpain activity was evident in both the cells in cytosolic and membrane fractions as measured using a fluorescent calpain substrate (Fig. 6A). Pretreatment of cells with the calpain inhibitor calpeptin was found to inhibit the activity of the calpains induced by PrP-(106-126) but not cell death. However, when cells were pretreated with a combination of 100 µM z-VAD and 100 µM calpeptin, apoptosis induced by 100 µM PrP-(106-126) was significantly inhibited (Fig. 6B).


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  		Fig. 6.   Increased calcium levels lead to calpain activation, which can be blocked by calpeptin. A, an increase in activity of the calcium activated calpain proteases, as measured by a fluorescent calpain substrate, is evident within 15 min of treatment with 100 µM PrP-(106-126). Activity was increased in both the cytosolic (black-square) and membrane () fractions. This activity was inhibited using 100 µM calpeptin. Column 1, untreated control; column 2, 100 µM calpeptin; column 3, 100 µM PrP-(106-126); column 4, 100 µM PrP-(106-126) + 100 µM calpeptin; column 5, 100 µM PrP-(106-126)scr. B, calpeptin alone does not stop the cells from undergoing apoptosis, but a combination of z-VAD and calpeptin shows significant inhibition of apoptosis. Column 1, untreated control; column 2, 100 µM PrP-(106-126); column 3, 100 µM calpeptin +100 µM PrP-(106-126); column 4, 100 µM z-VAD + 100 µM PrP-(106-126); column 5, 100 µM z-VAD + 100 µM calpeptin + 100 µM PrP-(106-126).


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

The toxicity of the PrP-(106-126) peptide in a human neuronal cell line has not been demonstrated previously. Host expression of PrPC has been illustrated to be a prerequisite of PrP-(106-126) toxicity (14). Accordingly we observed the presence of PrPC in the SH-SY5Y cell line upon Western blot analysis (data not shown). In some instances the neurotoxicity of PrP-(106-126) has been found to depend on the co-culturing of neuronal cells with microglia (14, 35). However, PrP-(106-126)-induced apoptosis in SH-SY5Y cells does not appear to require microglia. Therefore, while the peptide may act through effects associated with the microglia to induce or enhance neuronal death in some instances, this is not exclusively the case as in this study it was found to act only upon the neuronal cells.

The physical nature of PrP-(106-126) is an important factor in its toxicity. It has been demonstrated that the toxicity of PrP-(106-126) is related to its ability to form aggregates (13, 24). Yet other fibrillar peptides (PrP-(106-114) and PrP-(127-147)) were not toxic, indicating that fibrillogenic properties alone are insufficient for toxicity. We examined the aggregation status of PrP-(106-126) when prepared in PBS and aged over a period of 8 days and found the peptide immediately aggregated on day 0 and increased its level of aggregation with time. This is consistent with a recent study showing that solutions of PrP-(106-126) prepared in PBS at concentrations higher than 40 µM have high beta -sheet content and contain macromolecular assemblies (36). The scrambled version of the peptide, PrP-(106-126)scr, does not show the same tendency to aggregate. This is reflected in our observation that PrP-(106-126) induces apoptosis as measured by the apoptotic hallmarks of phosphatidylserine flipping and DNA cleavage, whereas PrP-(106-126)scr does not.

Mitochondrial dysfunction is a well documented event in apoptosis (37, 38). A process known as permeability transition (PT) appears to be responsible for the loss of Psi m, leading to the opening of the PT pore and release of solutes from the mitochondria (39-41). Among the proteins released are apoptosis-inducing factor and cytochrome c. Release of these proteins leads to activation of caspases, a family of serine threonine proteases, and subsequently apoptosis (42). The present study has shown that PrP-(106-126) induces a decrease in mitochondrial membrane potential, leading to the release of cytochrome c into the cytosol and the activation of caspase-3.

Among the caspase targets cleaved in this system is Bcl-2. The Bcl-2 family of proteins encodes both positive (Bax, Bcl-Xs, Bad, and Bak) and negative (Bcl-2, Bcl-w, and Bcl-XL) regulators of apoptosis, whose primary site of action appears to be at the mitochondrion. Furthermore, it appears that relative ratios and interactions between family members is a key factor in deciding the fate of a cell (43). Bcl-2 is an antiapoptotic protein capable of inhibiting apoptosis induced by a wide variety of apoptotic stimuli in a broad range of cell types (44). PrP-(106-126) has previously been reported to have an effect on Bcl-2 levels in primary rat cortical cultures (45), and the amyloid-beta peptide of Alzheimer's disease, similar in some respects to PrP-(106-126), has been shown to cause a 50% decrease in the level of Bcl-2 protein in primary human neuron cultures within 6 h (46).

We observed that Bcl-2 becomes completely degraded in SH-SY5Y cells within 15 min of treatment with PrP-(106-126), whereas the levels of Bax (proapoptotic) and Bcl-XL (antiapoptotic) were not seen to change. This is consistent with the fact that a caspase-3 cleavage site is present in the loop domain of Bcl-2 (29). Furthermore, z-VAD, a pan caspase inhibitor, was found to block the depletion of Bcl-2. Such a cleavage would mean that Bcl-2 is losing its BH4 domain, which is essential to its antiapoptotic activity (47). The resulting 23-kDa C-terminal Bcl-2 fragment has also been shown to have proapoptotic activity (48). We were unable to detect breakdown products of Bcl-2 even when a polyclonal antibody was used (data not shown). This is possibly due to another factor bound to Bcl-2 that is obscuring the epitope sites. PrP has been found to directly associate with the C terminus of Bcl-2 in the yeast two hybrid system (49). Perhaps PrP-(106-126) is also capable of binding to Bcl-2 or of inducing cellular PrP to bind to it.

Despite its inhibition of Bcl-2 degradation, z-VAD was not found to inhibit mitochondrial depolarization or apoptosis. This illustrates that depolarization of the mitochondria occurs upstream of caspase activation and Bcl-2 cleavage, again highlighting the mitochondria as a key target for PrP-(106-126). But this also indicates that although caspases are activated by PrP-(106-126) to assist in apoptosis, it would appear that their activation is not essential for completion of the cell death program. This is not to be unexpected as there is now much evidence for the existence of caspase-independent apoptosis (39, 50-52). A recent study showing that caspase-3 activation by beta -amyloid and prion proteins is independent from their neurotoxic effects supports this conclusion (53).

If caspases are not essential to apoptosis in this system, then another factor is capable of mediating the cell death program. We first examined the possible role of oxidative species, which are known to be mediators of apoptosis in a number of systems (30, 31), including some instances of neuronal apoptosis (54). There have been many reports relating oxidative stress to cell death induced by PrP-(106-126) and PrPSC (55-57). However, we found no relationship between cell death induced by PrP-(106-126) and oxidative stress in this cell line.

The mitochondria are also involved in the regulation of calcium homeostasis (58). Recently it has been found that calcium homeostasis is disrupted in cerebellar granule cells of prion-deficient mice (59) and in the hypothalamic gonadotropin-releasing hormone neuronal cell line treated with PrP-(106-126) and beta -amyloid (60). PrP-(106-126) has also been found to exert effects on calcium homeostasis through impairment of L-type voltage-sensitive calcium channels (15, 61, 62). Calcium is also a known mediator of apoptosis in response to many stimuli (63). We observed a rapid and sustained increase in the cytosolic concentration of calcium in response to PrP-(106-126). Furthermore, we have eliminated the endoplasmic reticulum as a possible source for this calcium. The only other calcium store in the cell large enough to account for the rise in calcium levels observed is the mitochondria. The calcium chelator BAPTA-AM, which buffered the rise in calcium, did not, however, inhibit mitochondrial membrane depolarization. This could indicate that calcium release is upstream of PT, which leads to mitochondrial depolarization. PT is known to be induced by high intracellular calcium (58). However, it appears more likely that membrane depolarization and calcium release are occurring simultaneously as a result of PT as observed in calcium-treated rat liver and brain suspensions (64). Either way PT is triggered by high calcium concentrations, so an initial release of calcium will serve to further enhance PT, speeding up mitochondrial membrane depolarization and releasing more calcium. This could account for the speed at which both these events reach their peak.

As a result of the raised intracellular calcium, calpains were observed to increase in activity. Calpain I, a neutral calcium-activated protease, has previously been reported to be involved in neuronal apoptosis (65). Calcium also activates the phosphatase activity of a calcineurin-calmodulin complex, which dephosphorylates Bad, allowing it to translocate to the mitochondria where it plays a role in cytochrome c release (66).

Using the calpain inhibitor calpeptin, we attempted to rescue PrP-(106-126)-treated cells from apoptosis. But calpeptin was not sufficient to rescue the cells. However, when used in combination with z-VAD to block both caspase and calpain activity, there was a significant reduction in the levels of apoptosis observed. Hence PrP-(106-126) activates two pathways leading to apoptosis. One is governed by caspases, the other is governed by calpains. We envisage much cross-talk between these pathways under normal circumstances, but when one of the pathways is blocked the other is capable of completing the cell death program alone.

The initiation site of the apoptotic pathway in this system, whether involving caspases or calpains, is at the mitochondrion, and this appears to be the principal target of PrP-(106-126) toxicity. There is evidence in the literature supportive of a role for mitochondrial dysfunction in other neurodegenerative disorders such as Alzheimer's disease (67) and a form of Parkinson's disease (68). Studies of scrapie-infected hamsters have reported physical disruption of the mitochondria as well as decreased activity of important mitochondrial enzymes (69). PrP-(106-126) has been shown to be capable of uptake into cells in vitro (70), and just how this event occurs is currently under study. After uptake into the cell, the peptide may act directly on the mitochondria. One possible site of interaction is with Bcl-2, which is primarily located on the mitochondria. Bcl-2 has been found to bind PrPC in the yeast two-hybrid system (49). Evidence also exists for possible functional interactions between Bcl-2 and PrPC; as in PrP-null cells, which are vulnerable to apoptosis induced by serum withdrawal, Bcl-2 can protect against cell death (75). Perhaps an interaction between PrP-(106-126) and Bcl-2 interrupts an antiapoptotic activity of Bcl-2 and/or its interaction with native PrPC. PrP-(106-126) has also been found to form ion-permeable channels in planar lipid bilayer membranes (71). These channels are large enough and nonselective enough to mediate the discharge of membrane potential and allow passage of calcium ions. If PrP-(106-126) formed channels in the mitochondria, it could cause the observed mitochondrial membrane depolarization and calcium efflux leading to cell death. A similar disruption of calcium homeostasis through channel formation has been reported for Alzheimer's beta -amyloid peptide (72).

In conclusion, we have shown, for the first time, that PrP-(106-126) induces apoptosis in a human neuronal cell line. The initial site of action appears to be the mitochondrion where membrane depolarization and calcium release occurs. This is possibly through an interaction with the Bcl-2 protein, which is known to be present on the mitochondrion. Downstream of the mitochondrion the apoptotic program can be completed through the action of either caspases or calpains, and apoptosis can only be blocked by the inhibition of both.

    FOOTNOTES

* The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Dagger To whom correspondence should be addressed. Tel.: 353-21-4904068; Fax: 353-21-4904259; E-mail: t.cotter@ucc.ie.

Published, JBC Papers in Press, August 30, 2001, DOI 10.1074/jbc.M103894200

    ABBREVIATIONS

The abbreviations used are: PrPSC, scrapie isoform of prion protein; PrPC, native cellular prion protein; PT, permeability transition; Delta Psi m, change in mitochondrial membrane potential; scr, scrambled; PBS, phosphate-buffered saline; z-VAD-fmk, N-benzyloxycarbonyl-Val-Ala-Asp(O-Me) fluoromethyl ketone; MTT, 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide; PI, propidium iodide; JC-1, 5,5',6-6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolecarbocyanine iodide; FL-1, 530/30 band pass filter; FL-2, 585/42 band pass filter; BAPTA-AM, 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid tetrakis- (acetoxymethyl ester).

    REFERENCES
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
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